NMR Studies on Structure and Dynamics of the Monomeric Derivative of BS-RNase: New Insights for 3D...
Transcript of NMR Studies on Structure and Dynamics of the Monomeric Derivative of BS-RNase: New Insights for 3D...
NMR Studies on Structure and Dynamics of theMonomeric Derivative of BS-RNase: New Insights for 3DDomain SwappingRoberta Spadaccini1, Carmine Ercole2, Maria A. Gentile2, Domenico Sanfelice2, Rolf Boelens3, Rainer
Wechselberger3, Gyula Batta4, Andrea Bernini5, Neri Niccolai5, Delia Picone2*
1 Dipartimento di Scienze Biologiche ed Ambientali, Universita del Sannio, Benevento, Italy, 2 Dipartimento di Chimica, Universita degli Studi di Napoli ‘‘Federico II’’,
Napoli, Italy, 3 Department of NMR Spectroscopy, Bijvoet Center for Biomolecular Research, Utrecht University, Utrecht, The Netherlands, 4 Institute of Chemistry,
University of Debrecen, Debrecen, Hungary, 5 Dipartimento di Biotecnologie, Universita degli Studi di Siena, Siena, Italy
Abstract
Three-dimensional domain swapping is a common phenomenon in pancreatic-like ribonucleases. In the aggregated state,these proteins acquire new biological functions, including selective cytotoxicity against tumour cells. RNase A is able todislocate both N- and C-termini, but usually this process requires denaturing conditions. In contrast, bovine seminalribonuclease (BS-RNase), which is a homo-dimeric protein sharing 80% of sequence identity with RNase A, occurs natively asa mixture of swapped and unswapped isoforms. The presence of two disulfides bridging the subunits, indeed, ensures adimeric structure also to the unswapped molecule. In vitro, the two BS-RNase isoforms interconvert under physiologicalconditions. Since the tendency to swap is often related to the instability of the monomeric proteins, in these paper we haveanalysed in detail the stability in solution of the monomeric derivative of BS-RNase (mBS) by a combination of NMR studiesand Molecular Dynamics Simulations. The refinement of NMR structure and relaxation data indicate a close similarity withRNase A, without any evidence of aggregation or partial opening. The high compactness of mBS structure is confirmed alsoby H/D exchange, urea denaturation, and TEMPOL mapping of the protein surface. The present extensive structural anddynamic investigation of (monomeric) mBS did not show any experimental evidence that could explain the knowndifferences in swapping between BS-RNase and RNase A. Hence, we conclude that the swapping in BS-RNase must beinfluenced by the distinct features of the dimers, suggesting a prominent role for the interchain disulfide bridges.
Citation: Spadaccini R, Ercole C, Gentile MA, Sanfelice D, Boelens R, et al. (2012) NMR Studies on Structure and Dynamics of the Monomeric Derivative of BS-RNase: New Insights for 3D Domain Swapping. PLoS ONE 7(1): e29076. doi:10.1371/journal.pone.0029076
Editor: Piero Andrea Temussi, Universita di Napoli Federico II, Italy
Received August 12, 2011; Accepted November 20, 2011; Published January 12, 2012
Copyright: � 2012 Spadaccini et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permitsunrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by grants OTKA (Hungarian Scientific Research Fund) CK-77515 and TAMOP (Operative Program for the Renewal of theSociety) 4.2.1/B-09/1/KONV- 2010-0007 (GB) (Convergence) and 20088L3BP3_001 (DP). The funders had no role in study design, data collection and analysis,decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: [email protected]
Introduction
Protein subunits may interchange part of their polypeptide
chain, giving rise to inter-twinned dimers, or even higher order
aggregated forms. This phenomenon, also known as 3D domain
swapping, is often accompanied by additional biological activities
with respect to the parent monomeric protein, leading to new
functions or even to toxic effects associated to deposition diseases
[1]. To date, neither the molecular mechanism of 3D domain
swapping or its predictability have been elucidated (for a recent
review, see [2]).
For historical reasons [3], but mainly for the variety of
oligomers formed, bovine pancreatic ribonuclease (the well-known
RNase A) is considered a prototype of 3D domain swapped
proteins: dimers, trimers, tetramers or even higher order oligomers
of this protein have been extensively characterized, from structural
and functional point of view [4]. This protein provided also the
first experimental evidence that a single polypeptide chain may
dislocate both N- and C-termini [5,6], even in the same oligomeric
structure [7]. The capability to dislocate N-termini is a feature of
other mammalian ribonucleases [8,9], which often in the
aggregated state acquire a selective cytotoxicity towards tumour
cells [10]. However, all the multimeric forms of RNase A and
RNase-like proteins for biophysical studies have been prepared in
vitro, under non-native conditions produced by modifying either
the environmental physico-chemical conditions [11], or the
protein sequence [12,13]. Recently RNase A has been shown to
form a minor population of dimers when it is produced in bovine
cells [14]. However, only BS-RNase, which shares 80% sequence
identity with RNase A and a common catalytic mechanism [15],
offers the possibility to study the swapping process in vitro under
mild physiological conditions. The swapping endows BS-RNase
with new biological properties, including selective cytotoxicity
towards tumour cells [10]. In contrast to RNase A and all other
pancreatic-like ribonucleases, which are monomeric in their native
state, BS-RNase is a dimer constituted by two identical subunits
linked through two disulfide bridges, as well as by non-covalent
interactions. The native enzyme is isolated from seminal vesicles or
bull semen [16] as a mixture of swapped and un-swapped covalent
dimers, indicated as MxM and M = M respectively. The molar
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ratio of the two isoforms MxM and M = M is 2:1 at thermal
equilibrium. The X-ray structures of the two isomers [17,18]
revealed only small differences in their tertiary structures, located
essentially around the 16–22 hinge region, which connects the
dislocating N-terminal helix to the protein body. In contrast, there
are no differences in the quaternary structures of swapped and
unswapped forms, because the inter-subunit interface (the so-
called ‘‘open-interface’’ in the 3D domain swapping terminology
[2]), pre-exists already in the un-swapped counterpart. This
feature reduces significantly the entropic penalty associated with
the oligomerization processes. Hence in the case of BS-RNase the
dislocation of the N-termini requires only a partial unfolding step
associated with a small enthalpic barrier. As a consequence, for
this protein the swapping is a physiological, equilibrium process
[19] and the molar ratio between swapped and un-swapped forms
depends on tiny structural differences that influence the energetic
balance. Upon selective reduction of the interchain disulphide
bridges, under mild conditions [20], the swapped form of BS-
RNase retains a dimeric structure stabilized by non-covalent
interactions, whereas the unswapped one is converted into a
monomeric derivative, indicated henceforth as mBS.
A comparison of the NMR structure of mBS and RNase A,
which share more than 80% of the aminoacid sequence, showed a
close similarity between the two proteins except for the 16–22
hinge region, which has higher flexibility in mBS [21]. According
to this result and other suggestions, [22,23,24,25,26], the hinge
region may play an important role in the control of the swapping.
Inspired by these observations, we have engineered the BS-RNase
sequence by substituting either all the different residues of the
hinge loop or only the central Pro at position 19, with the
corresponding residues of RNase A. Surprisingly, none of these
variants displayed significant differences in the hinge region
flexibility and swapping extent [19,27], and the biological activity
was only marginally affected [28]. A similar behaviour has been
observed also in other swapped proteins, leading to general
conclusion that ‘‘…based on mutagenesis studies, it is doubtful that
sequence features alone determine whether a protein will undergo
domain swapping’’ [2]. On the other hand, 3D domain swapping
has been often connected to the instability of the monomeric
subunits. Therefore, we decided to perform a more detailed
investigation of the structural and dynamical properties of
monomeric derivative of BS-RNase. The in depth analysis of the
conformational stability of mBS could help to decide if the special
features of the swapping dimer arise from an intrinsic property of
its subunits, or are rather a consequence of the pre-existence of a
dimeric structure. In this paper, we report the refinement of the
structure of mBS, together with a characterization of the protein
flexibility, chemical stability, surface accessibility and hydration by
a combination of NMR techniques and Molecular Dynamics
(MD) simulations.
Results
Structure refinementThe refined structure of mBS (pdb ID code 2lfj) was calculated
out of 2252 unambiguously assigned distance constraints. These
correspond to an average of 18 restraints per residue.
The resulting structures satisfied the experimental constraints
with small deviations from the idealized covalent geometry, 98%
of the backbone torsion angles for non-glycine residues being
within the allowed regions in the Ramachandram plot. The final
structural statistics are reported in Table 1 in comparison with
those of the formerly published structure (pdb entry 1QWQ) [21].
The increased number of distance constraints makes the newly
refined structures more defined as indicated from the global values
of RMSD of the backbone and the side chains, respectively 0.730
and 1.139 A. A bundle of the 10 structures with lower energies is
reported in Figure 1A. In more detail, the higher resolution
allowed to extend the third helix by one residue and to observe a
better ordered loop spanning residues 65–72, i.e. the loop that in
the natural enzyme is involved in the deamidation of Asn 67 [29].
Despite the overall higher number of distance constraints, the
hinge loop region (residues 16–22) remains among the most
flexible regions of the protein, as indicated from the low number of
inter-residual NOEs for this region and in agreement with
previous [21,27] and new relaxation data (see below).
Regarding the three residues of the catalytic triad, His 119 is
present in almost all the structures in one conformation, His 12 is
present in two subsets of conformations whereas Lys 41 is very
disordered. This result represents a difference with respect to the
published crystal structure of the monomer, where the lysine side-
chain is bound to a phosphate ion, whereas our solution is
deprived of any salt.
Hydrogen exchange measurementsFurther information on mBS stability were derived from
hydrogen exchange experiments on 15N labeled mBS at
pH 5.65, collecting a series of 1H-15N HSQC experiments at
300 K upon dissolving in D2O a lyophilized protein sample.
Table 1. Experimental restraints and structural statistics forten lowest-energy NMR structures of mBS-RNase (2lfj).
2lfj 1QWQ
NMR distance constraints
Intra-residue, (i2j) = 0 1056 601
Short-range, (i2j) = 1 512 291
Medium-range, 1,(i2j),5 216 123
Long-range, (i2j)$5 456 260
Total distance restraints from NOEs 2240 1275
Total distance restraints from S-S bond 12 12
Total distance restraints 2252 1287
NMR dihedral restraints
W, Y 104 314
Structural statistics
RMS deviation for bond angles: 2.0u 1.9u
RMS deviation for bond lengths 0.010 A 0.010 A
Averaged pairwise RMSD (A)
Backbone (global) 0,7360,12 1.2060,55
All heavy (global) 1,1460,14 1.7360,20
Backbone (secondary structure)1 0,4560,17 0,7660,10
All heavy (secondary structure)1 0,9160,20 1,3260,44
Structure quality by procheck analysis1
Residues in most favoured regions 82.8% 72.4%
Residues in additional allowed regions 14.1% 23,9%
Residues in generously allowed regions 1.7% 2,5%
Residues in disallowed regions 1.4% 1,2%
For comparison, also the corresponding data relative to the previous calculatedstructure (1QWQ) are reported.1Residues 4–12,25–31,43–64,70–87,95–112,115–117,122–124.doi:10.1371/journal.pone.0029076.t001
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Exchange constants could be evaluated for 45 well resolved
residues distributed all over the sequence, while the H/D exchange
process of most of the other amide groups have been only
qualitatively determined, see Table 2. Measured exchange constants
differ by more than three orders of magnitude along the sequence and
were slower by 3 orders of magnitude compared to the intrinsic rates.
For an evaluation of kop protection factors and DG based on the
exchange rates, it is important to elucidate before the mechanism
by which exchange occurs in the conditions employed for the
kinetic measurements [30]. It has been reported that under
unimolecular exchange regime (EX1) there is no pH dependence
of Kex, whereas in the bimolecular exchange limit (EX2) Kex will
be proportional to the pH [31]. Accordingly, to provide insight
into the exchange mechanism we measured the exchange rates
also at pH 6.7. The data obtained indicated for BS-RNase an EX2
mechanism, as in the case of RNase A [32].
Measured exchange rates at pH 5.65 were used to calculate
protection factors and DGop, which are reported in Table 2 and
plotted in Figure 1B. All the regions of regular secondary structure
are well protected from the solvent. As a further evidence of the
extreme stability of mBS structure, it’s interesting to notice that
also residues of some loop regions, such as residues 67–69
belonging to the deamidation loop, exchange slowly.
Relaxation measurementsConventional 15N relaxation data (T1,T2, NOE), 15N/1H CSA/
DD cross-correlated cross-relaxation rates and heteronuclear
NOEs, revealed that residues 2, 18, 20, 21, 22 and 69, 70, 71,
72 were outliers. These data confirm the earlier observations [21],
that predicted a high flexibility of the 16–22 hinge and 65–72 loop
regions. Residue 2, on the other hand, belongs to the disordered,
solvent exposed N-terminal extremity, hence the outstanding
values of its relaxation parameters. Independently of the involve-
ment of the R2/R1 outliers, the model-free evaluation [33] of the
T1, T2 and NOE data set yields tc = 7.9 ns global correlation time
and an average of S2 = 0.9060.07 for the order parameters
(overlapping signals and exchange affected residues 70–72 were
intentionally omitted from the fit). Similarly to raw relaxation data,
S2 order parameters also exhibited dips in the hinge region around
S18 (Figure 2). The back-calculated ‘theoretical’ T1, NOE and T2
relaxation data derived from these parameters agreed with the
experimental values within 5, 6 and 9% (1 standard deviation),
respectively. This modest agreement may be a sign of more
mobility, since the typical experimental accuracy of the exponen-
tial T1 and T2 fits was within 2%. Visualisation with reduced
spectral density mapping (Figure S1) [34] is a useful complement
of the model-free method. On the J(vN)/J(0) spectral density
correlation map we see that the bulk of the residues are below the
descending part of the limiting curve of the rigid body approach,
due to limited internal motions. However, residues that are
affected by exchange are right shifted because of increased J(0)
contribution (residues 70, 71 and 72). Both approaches gave a
dominant monomeric structure, with a correlation time appropri-
ate for this size of proteins. Indeed, a simple empirical formula
[35] predicts 8.5 ns correlation time at 300 K for a 124 residue
globular protein in water, that is in reasonable agreement with our
Figure 1. NMR derived structure of mBS. A) Bundle of the best 10 structures of mBS calculated out of 2252 distance constraints (pdb ID code2lfj); B) protection factors mapped on mBS structure with the following colour code: residues with a P.16105 in red, the one with 106103,P,16105
in orange and the one with P,106103 in yellow. Residues that couldn’t be analyzed are shown in grey.doi:10.1371/journal.pone.0029076.g001
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findings. The small difference might be a consequence of
anisotropic rotational diffusion. 15N transversal CSA/DD cross-
correlated cross-relaxation rates again displays the same mobile
regions of mBS (see Figure S1). Combination [36] of 15N CSA/
DD cross-correlated and conventional relaxation data however,
predicted an unlikely high exchange contribution to R2 relaxation
over the whole sequence. Overall, the relaxation approach
described here unequivocally identified two mobile regions in
mBS, the hinge 16–22 and the loop 69–72 region, in good
agreement with previous heteronuclear NOE data of the same
protein and of RNase A. However, careful inspection of the NOE
(Figure S2) and S2 data anticipates enhanced dynamics of the 113–
115 turn region as well. Moreover, extra mobility can persist in the
119–121 region of the b-sheet, as it can be seen from reduced
spectral density mapping and R2/R1 rates (Figure S2). This seems
to support the curling of the 115–123 strand in the ensemble of the
refined NMR structures.
Molecular Dynamics simulationOn the basis of the refined mBS solution structure (pdb ID 2lfj) a
100 ns MD run has been performed in order to predict i) flexibility
along the protein backbone, ii) stability of intramolecular hydrogen
bonding network and iii) modes of the interaction between mBS
and water molecules. In Figure 3 rmsf values are reported for all
backbone amides, predicting the highest flexibility for the N
terminal moiety. Flexibility is also predicted for 65–68 and 111–113
fragments whose rmsf values are well above the standard deviation,
s= 1.2 A, calculated for all the residues outside the N terminus.
Surprisingly, an average rmsf value of 2.2, just above s, is observed
for the 16–22 hinge loop, that in all the experimental measurements
showed a significant flexibility. By analysing the MD trajectory,
hydrogen bond percentage lifetimes have been estimated for all
mBS backbone amide hydrogens according to previously reported
procedures [37,38]. The obtained results, compared also with the
corresponding atom depth indexes, are shown in Table 1S.
Reduced involvement in intramolecular hydrogen bonding, as
expected, is predicted only for outer backbone amide hydrogens.
Density of water molecules in contact with mBS has been derived
from the MD trajectory and the distribution of MD hydration sites,
MDHS, is also reported in Table 1S.
Urea unfoldingTo characterize the unfolding of mBS we have studied its
denaturation in a residue specific manner collecting a series of1H-15N HSQC NMR spectra at increasing urea concentration,
ranging from 1 to 7 M, and monitoring peak volumes. Figure S3
shows the spectra measured in the presence of 0, 1, 3 and 6 M
urea respectively. The HSQC spectra undergo minor chemical
shift changes at increasing urea concentration. At low urea
concentration (1 M) most of the residues of the monomeric
proteins are unaffected from the presence of the denaturant, only
the residues of the central strands show a chemical shift
perturbation higher than 0.1 ppm. However, by increasing the
urea concentration up to 3 M, peaks originating from denatured
state appear. As the urea denaturation was found completely
reversible, the simultaneous presence of peaks belonging to the
folded and unfolded state indicates slow conformational exchange
on the NMR chemical shift timescale.
The intensities of 30 residues distributed along all the sequence,
shown in Figure 4A, could be followed unambiguously up to 6 M
urea. This data indicates that still at high urea concentration there
is some localized structure. In figure 4B are shown the changes in
the normalized NMR peak intensities for selected residues across
the range of urea concentrations studied.
Table 2. Summary of Kex. protection factor (P) andDGopderived from hydrogen exchange experiments at pH 5.7.
Residue Kex(hr21)a P DGop(kcal mol21)
PHE8 0.11460.046 8.87E+03 5.4060.04
GLU9 0.27360.015 1.82E+03 4.4660.02
ARG10 0.13060.018 9.03E+03 5.4160.06
GLN11 0.02660.015 9.75E+04 6.8260.04
HIS12 0.03860.016 2.48E+05 7.3860.06
ASP14 0.19060.027 4.83E+03 5.0460.01
SER21 0.04060.100 1.56E+05 7.0160.03
CYS26 0.10960.057 4.86E+04 6.4160.02
LEU28 0.64060.073 1.15E+03 4.1860.02
MET30 0.01160.016 1.53E+05 7.0960.02
CYS31 0.13460.019 4.54E+04 6.3760.01
CYS32 2.22060.570 6.13E+03 5.1860.02
ARG33 0.24960.028 1.85E+04 5.8460.01
LYS34 0.26260.017 7.70E+03 5.3260.05
LYS41 0.20060.084 1.75E+04 5.8060.04
VAL43 0.46960.101 3.39E+02 3.4660.06
ASN44 0.03160.010 9.62E+04 6.8160.09
GLU49 0.04460.015 6.78E+04 6.6160.01
ASP53 0.72560.120 9.90E+02 4.1060.03
ALA56 0.05960.006 2.96E+04 6.1260.02
VAL57 0.00360.007 7.90E+04 6.7060.04
CYS58 0.01060.006 3.39E+05 7.5660.01
SER59 0.42560.052 2.12E+04 5.9160.01
LYS61 0.06260.075 3.11E+04 6.1460.02
GLN69 0.02660.072 8.70E+04 6.7660.04
GLN74 0.00560.014 3.43E+05 7.5760.05
SER77 0.01160.044 3.73E+05 7.6260.03
ARG80 0.19160.026 1.08E+04 5.5260.02
ILE81 0.00760.015 5.91E+04 6.5360.06
THR82 0.00560.021 1.34E+05 7.0160.05
ASP83 0.18260.025 6.19E+03 5.1960.03
CYS84 0.02760.017 1.21E+05 6.9560.04
ARG85 0.01660.015 2.88E+05 7.4760.01
GLU86 0.08360.016 8.63E+03 5.3860.09
SER89 0.03360.025 1.40E+05 7.0460.03
LYS91 0.03860.013 6.38E+04 6.5760.10
ALA96 0.62160.112 6.21E+03 5.1960.08
TYR97 0.25260.034 2.84E+03 4.7360.02
THR100 0.05760.015 3.15E+04 6.1560.06
VAL102 0.02360.010 1.85E+04 5.8360.02
LYS104 0.01460.079 5.44E+04 6.4860.04
ILE106 0.00860.013 1.61E+05 7.1260.02
ALA109 0.01460.014 6.89E+04 6.6260.07
GLY111 0.32260.086 2.21E+04 5.9460.06
VAL118 0.01260.022 1.30E+04 5.6360.02
HIS119 0.02660.012 1.66E+05 7.1460.05
VAL124 0.36060.038 3.26E+01 2.0760.04
aReported errors refer to the errors in the least square fitting.doi:10.1371/journal.pone.0029076.t002
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A two state mechanism of unfolding, in which only native and
denatured molecules are considered [39], fits well with the
experimental NMR data. From the data fitting we could calculate
the concentration of urea at midpoint of unfolding (C1/2). The
selected residues show very similar values of C1/2, indicating that
all the mBS regions behave similarly as a function of urea
concentration. The average concentration of urea at midpoint of
unfolding is 4.2 M, a value very close to the one obtained
monitoring the secondary structure changes at increasing urea
concentration by far-UV CD [40]. This result definitely confirms
also that the unfolding of mBS follows a two state mechanism,
without the presence of intermediate states.
Paramagnetic profile of TEMPOL accessibility to mBSsurface
TEMPOL, the soluble and stable free radical commonly
employed for analyzing the distribution of protein surface hot
spots [41] has been used as paramagnetic probe for investigating
the surface accessibility of mBS. Changes of 1H-15N signal
intensities of backbone amides in HSQC protein spectra,
recorded in the presence of variable amounts of TEMPOL in
solution, have been measured and reported as paramagnetic
attenuations, Ai.
As shown in Table S1 Ai have been calculated for most of mBS
amide groups, i.e. 87 out of the total 118 well resolved NH signals
which are present in diamagnetic and paramagnetic 1H–15N
HSQC spectra. The obtained Ai values range from a maximum of
2.0 to a minimum of 0.4 respectively for signals exhibiting strong
and weak paramagnetic attenuations. M13, S15, T45, C65 and
G112 residues, exhibiting Ai values higher than 1.7, are the most
attenuated amide signals. Interestingly, MD simulation results (vide
supra) predict that all of them are not tightly involved in
intramolecular hydrogen bonding, see Figure 5 and Figure S4,
confirming the already suggested bias of TEMPOL to interact
with accessible backbone hydrogen bond donors [42].
Figure 2. Relaxation data of mBS. S2 order parameters of mBS as obtained from the model-free analysis of Lipari and Szabo. (Details are given inthe text.) Error bars within the 90% confidence limit were calculated with Montecarlo method Outliers according to R2/R1 rates (residues 70–72) wereintentionally omitted from the simultaneous fit of the global correlation time, effective correlation times and order parameters. The mobility of theseresidues is better demonstrated with the reduced spectral density mapping (see supplementary material).doi:10.1371/journal.pone.0029076.g002
Figure 3. Theoretical Molecular Dynamics of mBS. Root mean square fluctuations (rmsf) values vs. mBS sequence.doi:10.1371/journal.pone.0029076.g003
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Discussion
Although 3D domain swapping was proposed about 50 years
ago [3], and it is actually well represented in PDB, its molecular
mechanism is still unknown. Eisenberg et al. provided its first
definition and proposed a general scheme for this process. In the
simplest case ‘‘closed’’ monomeric proteins convert themselves in
‘‘open’’ monomers through a local unfolding step. Then, the
interaction between two or more open monomers gives rise to
swapped dimers, or even higher oligomers [43]. Very recently,
Laurents and coworkers [11] proposed a different mechanism,
which involves the transition through an ‘‘activated’’, unfolded
state, re-opening the debate and the scientific interest on the 3D-
domain swapping. With respect to the proteins considered by both
authors, however, BS-RNase presents still some singularities,
which are worth of attention. Indeed, interchange of N-terminal
extremities is a physiological, equilibrium process which does not
require severe environmental conditions. The two subunits of BS-
RNase are covalently linked through two disulfide bonds, so that
the two subunits are already correctly oriented for the swapping.
In vivo, in the reducing cytosol compartment, the un-swapped
isomer is supposed to be converted into two monomers, whereas
the swapped isoform, stabilized by the interchanged N-terminal
helices, retains a dimeric structure [20]. Therefore, BS-RNase is
supposed to occur in vivo under multiple forms, depending on the
biological compartment encountered, with different tertiary and/
or quaternary structures. Given the importance of the 3D domain
swapping in the biological properties of BS-RNase, in this paper
we have performed a deeper investigation of the structural and
dynamical features in solution of its monomeric derivative, with
the aim to understand if the BS-RNase subunit has an intrinsic
feature to undergo a local unfolding and to be converted into an
open monomer, according to the Eisenberg hypothesis [44]. As a
first step, we report here the refinement of the solution structure by
Figure 4. Urea perturbation of mBS structure. A) On mBS structure are shown in red residues whose intensities were still detectable at 6 Murea; B) changes in the normalized NMR peak intensities for selected residues across the range of urea concentration studied. The line/symbol codefor each residue is the following:& K39, ¤ S77, m L32, n K26, * H12, m G111, dashed line V118, & V54, dashed-dotted line G88, ¤ D83, # E49, %- -T36.doi:10.1371/journal.pone.0029076.g004
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high resolution NMR spectroscopy up to 0.730 and 1.139 A
resolution for backbone and all atoms respectively, combined with
relaxation measurements in solution at 500 MHz and with a
characterization of the surface accessibility and hydration at
atomic resolution. The increased number of restraints used for
structure calculation allowed us a better definition not only of the
secondary structure elements, -resulting in a lengthening of helix 3
by one residue- and of the side-chains, but also of the loops,
including the 65–72 region. This loop plays a remarkable role in
the biological properties of BS-RNase because it includes the Asn-
Gly sequence, which spontaneously deamidates under physiolog-
ical conditions [29], and represents also a potential contact region
with Ribonuclease Inhibitor [45], a horse-shoe shaped protein
which readily and efficiently inactivates most pancreatic-like
ribonucleases. Structural and relaxation data indicate that the
mBS behaves like a compact, globular monomer, and the
correlation time allowed us to exclude not only the presence of
dimeric forms, which instead have been observed in the
homologue human pancreatic protein [46], but also a partial
opening of the protein. Moreover relaxation data highlight the
presence of two main flexible regions, corresponding to the 16–22
and 65–72 loops; in addition, extra-mobility is found at the C-
terminal region (112–115 loop and 119–121 strand), i.e. in the
region involved in the swapping of the C-terminal strand of RNase
A [6]. A cis-trans isomerisation of proline at position 114 is involved
in this process but, despite the presence of a cis Pro in the same
position, the swapping of the C-terminal ends has never been
observed in BS-RNase.
The preferential access of TEMPOL towards few backbone
amide groups yields relevant information on mBS surface
accessibility. Indeed, the presence of backbone amide groups
which are, at the same time, surface exposed, available for
hydrogen bonding and TEMPOL accessible, as in the case of
M13, S15, C65 and G112 residues, is diagnostic of surface hot
spots, belonging to the N-terminal arm (M13, S15), the
deamidation loop (C65) and the C-terminal hinge loop (G112).
On the other hand, the high paramagnetic attenuation of T45
amide signal is consistent with the expected enhanced accessibility
of active site regions [18]. Combined analysis of data obtained
from H/D exchange and paramagnetic perturbations indicates
that amide groups of residues exhibiting the highest Ai values are
also involved in fast H/D isotopic exchange. However, fast
isotopic exchange rates do not imply strong paramagnetic
perturbations. In this respect, Y92 NH group represents a limiting
case, as it exhibits, at the same time, very fast exchange rate and
the lowest Ai value. The presence of a MDHS close to Y92 NH
could account for local presence of structured water molecules
which prevent free TEMPOL approach. It is worth noting that
high TEMPOL accessibility of S15 and C65 residues has been
already observed in the corresponding positions of the homologous
bovine RNase A in a recent multiple solvent crystal structure
(MSCS) investigation. In the latter protein, indeed, S15 and C65
residues were in close proximity of R,S,R-bisfuran alcohol and
dimethylsulfoxide respectively [47], suggesting some binding
properties of these enzyme moieties also in solution.
Thus, high compactness of the protein structure is consistently
suggested by the analysis of the H/D exchange data and
TEMPOL surface accessibility profiles, as secondary structure
elements are all protected from the solvent. This result is very
interesting and somehow unexpected, if we consider that helix 1
(residues 4–12) corresponds to the region swapping between the
two isoforms, and confirms a close similarity with RNase A
structure. In addition, the monitoring of urea denaturation of mBS
by NMR allowed us to follow the unfolding process in a residue
specific way and to elucidate, eventually, the presence of residues
more prone to unfolding and possibly prompting dislocation and
swapping. The analysis reported in figure 4 gives us a convincing
evidence of a two-step denaturation mechanism for all the protein
regions, in agreement with a recent CD and calorimetric study
[28], showing that the protein denaturation follows a two step
mechanism, and excluding the presence of significantly populated
intermediates. On the whole, all data presented in this paper
definitely confirm a close similarity with RNase A structure,
suggesting also that this protein is extremely compact and there is
no evidence of a pre-opening of the monomeric structure in
solution. With respect to our previous data [21], this more detailed
Figure 5. Tempol surface mapping of mBS. Paramagnetic attenuations, Ai, are reported for each well resolved 1H-15N HSQC signal of mBS (filledtriangles). Histogram heights refer to the fractional freedom from intramolecular hydrogen bonding predicted by the 100 ns MD simulation in explicitwater (grey bars).doi:10.1371/journal.pone.0029076.g005
Structure and Dynamics of Monomeric BS-RNase
PLoS ONE | www.plosone.org 7 January 2012 | Volume 7 | Issue 1 | e29076
investigation highlights the presence of potential hot-spots along
the protein structure, in particular in the C-terminal region,
henceforth we cannot exclude the possibility that, in different and
eventually more severe conditions, also the C-terminal swapping
could be observed. Regarding the N-terminal hinge region, this is
still one of the most flexible regions of the protein but, according to
previous mutagenesis studies [19,48], hinge flexibility together
with swapping propensity and in vitro antitumor activity [28], are
not strictly dependent on a specific sequence.
On the other hand, we cannot exclude other structural and
functional roles for the 16–22 loop, which shows a very low
similarity with the corresponding region of RNAse A. It is worth to
recall here that the presence of a proline replacing Ala 19, in the
middle of this region, prevents the subtilisin cleavage [49], and
inactivation observed in RNase [50], so that we may hypothesize
that the 16–22 hinge sequence of BS-RNase makes the protein
more resistant to protease attack. It is also worth to recall here that
in a very recent paper by Eisenberg and coworkers [51] the region
encompassing Ala 19 in RNase A has been identified as a very
good candidate to prompt protein aggregation, thus the possibility
that the substitutions present in the hinge region of BS-RNase [52]
have been selected to prevent self-assembly phenomena deserves
also consideration. For all these reason we believe that the unusual
properties of the swapping process of BS-RNase are not strictly
dependent on its N-terminal hinge region, as initially supposed,
but on the pre-existence of a dimeric structure, independently on
the swapping. With respect to RNase A the two extra disulphides
at position 31 and 32, that convert the protein into a dimer, on one
end facilitate the N-swapping, reducing the entropic disadvantage
associated to other 3D domain-swapping cases, all consisting in a
momomer/swapped dimer conversion and also preparing the two
subunits in the correct orientation, and on the other end create a
steric encumbrance which hinders the C-swapping. Finally, the
possibility that very subtle structural and dynamical differences
induced by inter-subunits interactions increase the protein
plasticity should also been considered. Further mutagenesis
studies, together with the on-going characterization of the dimers
in solution will be helpful to confirm this hypothesis and to
understand the molecular basis of the swapping process in BS-
RNase.
Materials and Methods
NMR experiments15N and 13C,15N labelled monomeric BS-RNase was expressed
and purified as previously described [21]. NMR spectra for
structure refinement were acquired at 300 K on Bruker spec-
trometers operating at 500, 600, 700 and 750 MHz, equipped
with triple resonance gradient probes. Data were processed with
NMRPipe [53]; visualization of spectra, peak-picking and analysis
were done in NMRView [54]. Chemical-shift assignments were
obtained from standard three-dimensional triple resonance
experiments recorded on 15N,13C-labeled samples of mBS [55].
NOE-based distance restraints were extracted from a 500-MHz
2D NOESY spectrum, a 500-MHz 3D NOESY-[1H,15N]-HSQC
spectrum, and a 700-MHz 3D NOESY-[1H,13C]-HSQC spec-
trum, all with a mixing time of 100 ms. 208 dihedral angle
constraints were derived from chemical shifts values using TALOS
[56]; additional restraints were obtained from 47 3JHNHA coupling
constants extracted from a 500-MHz quantitative J-correlated
HNHA experiment [57].
The urea induced unfolding of BS-RNase was performed
adding solid urea (uread4, Sigma Aldrich) directly into the NMR
tube. The urea concentration was increased by 1 M at each
titration step until 7 M. In in order to keep protein concentration
and pH constant we prepared a different protein sample for each
point of the titration and for each of them we adjusted the pH at
5.7. To check for the possible volume increase caused by the
addition of solid denaturant, the length of the NMR sample was
measured at each step and found constant (within the experimen-
tal error). The unfolding process was monitored with a series of 2D
HSQC experiments collected at 300 K.
Structure calculationStructure calculations were performed by simulated annealing
in torsion angle space, using the CYANA 2.0 package [58], which
implements an efficient protocol for structure calculation/
automated assignment of NOEs. The standard annealing protocol
was used with 20000 steps of torsion angle dynamics; in each of
seven cycles, 100 structures were calculated, and the 20 with the
lowest target function were used in the next stage. In the final run,
200 structures were computed, and the 40 with lowest target
function were refined by 4000 steps of restrained minimization in
the more realistic AMBER99 force field [59]. To mimic the effect
of solvent screening, all net charges were reduced to 20% of their
real value, and a distance-dependent dielectric constant (e= r) was
used. The cutoff for non-bonded interactions was 12 A. The
NMR-derived upper bounds were imposed as semi-parabolic
penalty functions, with force constants of 16 kcal mol21 A22; the
function was shifted to linearity when the violation exceeded
0.5 A.
The AMBER-minimized structures were sorted by increasing
restraint violation, and the first 10 were selected as the
representative bundle. Quality checks were done in PRO-
CHECK-NMR [60], while MOLMOL [61] was used for
structure visualization and analysis.
Molecular Dynamics simulationsMD simulations were performed in explicit solvent starting from
the lowest energy NMR structure of (pdb ID 2lfj). GROMACS
package [62] and the AMBER force field [63] were used for the
100 ns MD run of solvated structure in a cubic box of equilibrated
TIP3P water molecules [64]. The initial shortest distance between
the protein and the box boundaries was set to 1.0 nm and chloride
ions were added to achieve global electric neutrality. Afterwards,
the system was energy minimized with 900 steps of conjugate
gradients. To achieve a good equilibration prior to the long MD
simulations, the system was subjected to a short (20.0 ps) MD run
where the atoms of the macromolecule were restrained to their
positions, allowing only the solvent to move. The protein-water
system was simulated in the NPT ensemble by keeping constant
the temperature (300 K) and pressure (1 atm); a weak coupling to
external heat and pressure baths was applied (relaxation times
were 0.1 ps and 0.5 ps, respectively). Bonds were constrained by
LINCS algorithm, while non-bonded interactions were accounted
by using the PME method (grid spacing 0.12 nm) [65] for
electrostatic contribution and 0.9 nm cut-off for VDW contribu-
tion. An integration time step of 2 fs was used and trajectory
snapshots were saved every 0.2 ps. As the backbone RMSD levels
off after an equilibration period of 100 ps subsequent analysis of
MD trajectories was carried out from 100 ps onward. Cut-off
values of 0.35 nm and 75u between donor and acceptor moieties
have been considered for hydrogen bonding. The fraction of the
MD trajectory in which hydrogen bond criteria are fulfilled for a
given donor is denoted by ‘‘hydrogen bond percentage life time’’
[37,38]. Solvent density map, whose maxima have been defined as
molecular dynamics hydration sites (MDHS’s) [66,67], were
calculated from the atomic coordinates of the explicit waters in
Structure and Dynamics of Monomeric BS-RNase
PLoS ONE | www.plosone.org 8 January 2012 | Volume 7 | Issue 1 | e29076
the simulation. The space surrounding the protein was divided
into two shells: the first one extended up to a distance of 0.6 nm
from the protein surface and accounted for the protein hydration
sites; the second one, representing the bulk solvent, extended from
0.6 nm to 0.8 nm from the protein surface. The water positions
were accounted in a 3D grid (step-size 0.05 nm). For each frame,
the protein was superimposed onto a reference structure in order
to eliminate the effects of translations and rotations. 3D iso-
contour plot of the resulting water density was obtained by using
PyMOL software (http://www.pymol.org).
Hydrogen exchangeHydrogen exchange experiments were performed dissolving the
15N-labelled lyophilised NMR protein sample in 99.9% D2O. A
series of successive 2D 1H-15N HSQC spectra was recorded and
300 K. Fifty hydrogen exchange rates were determined by fitting a
first order exponential equation to the peak heights versus time data:
I(t)~I(0) exp ({kobst)
where I (t) represents the cross peak height at time t, I(0) the original
cross peak height, kobs the observed rate of hydrogen exchange and t
the time in seconds. The intrinsic hydrogen exchange rates krc were
calculated using the web server program Sphere (http://www.fccc.
edu/research/labs/roder/sphere, Yu-Zhu Zhang, Protein and
peptide structure and interactions studied by hydrogen exchange
and NMR. Ph.D. Thesis, Structural Biology and Molecular
Biophysics, University of Pennsylvania, PA, USA.). The protection
factor (PF) for each well resolved NH group was then calculated
using the equation:
PF~krc=kobs
The free energy of opening, DGop, was calculated from the equation
DGop~{RTlnKop
where kop = kobs/krc. Data were fitted with the program Kaleida-
Graph 3.51 (Synergy software) [68].
RelaxationFor NMR dynamics, besides the conventional 15N relaxation
(T1, T2 and 15N-NOE) [69], the longitudinal and transverse 15N
and 1H CSA/DD cross-correlated relaxation rates gzz(15N),
gxy(15N), gxy(
1H) [46], were measured at 500.13 MHz on a
Bruker DRX spectrometer. To this end, a series of 2D 1H-15N
HSQC spectra using sensitivity enhanced gradient pulse schemes
[70] were recorded. The typical experimental parameters are
summarized as follows. The 1H carrier frequency was set to the
water resonance at 4.7 ppm using 15 ppm window, while the 15N
window was 28 ppm centred at 118.5 ppm. The relaxation delay
times were set as follows for T1: 11.2, 101.2, 201.2, 401.2, 601.2,
801.2, 1001.2, 1201.2 ms; for T2, CPMG pulse trains of 0.03,
30.4, 60.8, 91.2, 121.6, 182.4, 243.2, 302.4, 360, 417.6 ms in
duration were used; for the measurement of cross-correlated
relaxation rates, gzz and gxy, the relaxation interference was
allowed to be active for 20 or 21.6 ms in pairs of experiments
including the reference (D= 0 ms) experiment. The number of
transients collected per t1 increment was 8 for T1, 16 for T2, 32
for NOE, 128 for gzz and 80 for gxy measurements. A spin-lock
field of 3400 Hz was used for the 15N transverse cross-correlation
experiment. Two-parameter exponential fits of the measured
volume intensities of cross-peaks were applied to extract the
relaxation times T1 and T2. The cross-correlation rate constants
were determined using the initial linear build-up rate approach.
The theoretical expressions for the autorelaxation (R1, R2) and
cross-correlation rate constants (gxy, gzz) and for the steady-state
heteronuclear NOE in terms of the spectral density functions
(Ja(v) auto- and Jc(v) cross-correlation) are used as given in the
literature [71]. The simplifying assumption of isotropic overall
tumbling and the axial symmetry of constant (Ds= 2160 ppm)15N chemical shielding tensors were applied. A bond length of
rNH = 0.102 nm was used in all calculations. The model-free [33]
analysis of T1, T2 and heteronuclear NOE yielded the S2 order
parameters and local correlation times for most amides and the
global correlation time.
Paramagnetic studies of Tempol induced perturbations1H–15N HSQC diamagnetic and paramagnetic spectra, record-
ed from a Bruker DRX 600 spectrometer with 512 increments and
128 scans over 2048 data points, were compared to determine
paramagnetic perturbations on signal intensities.
Only well resolved NMR signals have been considered for
quantifying cross-peak volumes in the presence and in the absence
of the paramagnetic probes, respectively Vid and Vi
p. Such volumes
have been measured with an estimated error lower than 10% and
their autoscaled values, ui, were used according to the relation:
up,di ~
Vp,di
(1=n)P
n
Vp,di
Paramagnetic attenuations, Ai, were calculated from the auto-
scaled diamagnetic and paramagnetic peak volumes, respectively,
ud and up, according to the relation:
Ai~2{u
pi
udi
Atom depth calculationsBy using the SADIC suite [72], atom depth of all the hydrogen
of mBS backbone amide groups have been calculated. A radius of
8 A for the probing sphere was employed to measure exposed
volumes and, hence, depth indexes, Di.
Supporting Information
Table S1 Experimental and calculated parametersobtained for solvent-protein interactions.
(DOC)
Figure S1 Relaxation data of mBS. A) Residue by residue
representation of 15N spectral densities as a function of low-
frequency components. The continuous curve represent a rigid
body, with 8 ns global correlation time. The outliers to the right of
the curve clearly identify residues 70–72 as the most exchange
affected part of mBS; B) Transversal 15N-1H CSA-DD cross-
correlated cross-relaxation rates of the amides in mBS. Lower rates
can be attributed higher internal mobility.
(TIF)
Figure S2 Relaxation measurments on mBS. A) Hetero-
nuclear 15N-{1H} NOE values in mBS; B) Ratio of 15N T1/T2
relaxation times in mBS.
(TIF)
Structure and Dynamics of Monomeric BS-RNase
PLoS ONE | www.plosone.org 9 January 2012 | Volume 7 | Issue 1 | e29076
Figure S3 Urea denaturation experiments. 1H-15N
HSQC spectra of mBS at different urea concentrations.
(TIF)
Figure S4 Surface accessibility of mBS. Correlation
diagram between TEMPOL-induced paramagnetic perturbations
and fractional HB freedom predicted by MD simulation. The least
squares linear fit, represented by the straight line, yielded the
following parameters: slope 0.22, intercept 0.81, correlation
coefficient 0.18.
(TIF)
Acknowledgments
A special thank is due to Dr. Teodorico Tancredi for his help and constant
encouragement.
Author Contributions
Conceived and designed the experiments: DP RB RS NN GB. Performed
the experiments: RS CE AB RW GB DP. Analyzed the data: RS MAG DS
AB GB NN. Contributed reagents/materials/analysis tools: DP RB NN
GB. Wrote the paper: DP RS NN GB.
References
1. Bennett MJ, Sawaya MR, Eisenberg D (2006) Deposition diseases and 3D
domain swapping. Structure 14: 811–824.
2. Gronenborn AM (2009) Protein acrobatics in pairs–dimerization via domain
swapping. Curr Opin Struct Biol 19: 39–49.
3. Crestfield AM, Stein WH, Moore S (1962) On the aggregation of bovine
pancreatic ribonuclease. Arch Biochem Biophys Suppl 1: 217–222.
4. Libonati M, Gotte G (2004) Oligomerization of bovine ribonuclease A:
structural and functional features of its multimers. Biochem J 380: 311–327.
5. Liu Y, Hart PJ, Schlunegger MP, Eisenberg D (1998) The crystal structure of a
3D domain-swapped dimer of RNase A at a 2.1-A resolution. Proc Natl Acad
Sci U S A 95: 3437–3442.
6. Liu Y, Gotte G, Libonati M, Eisenberg D (2001) A domain-swapped RNase A
dimer with implications for amyloid formation. Nat Struct Biol 8: 211–214.
7. Liu Y, Gotte G, Libonati M, Eisenberg D (2002) Structures of the two 3D
domain-swapped RNase A trimers. Protein Sci 11: 371–380.
8. Canals A, Pous J, Guasch A, Benito A, Ribo M, et al. (2001) The structure of an
engineered domain-swapped ribonuclease dimer and its implications for the
evolution of proteins toward oligomerization. Structure 9: 967–976.
9. Merlino A, Avella G, Di Gaetano S, Arciello A, Piccoli R, et al. (2009) Structural
features for the mechanism of antitumor action of a dimeric human pancreatic
ribonuclease variant. Protein Sci 18: 50–57.
10. Kim JS, Soucek J, Matousek J, Raines RT (1995) Catalytic activity of bovine
seminal ribonuclease is essential for its immunosuppressive and other biological
activities. Biochem J 308(Pt 2): 547–550.
11. Lopez-Alonso JP, Bruix M, Font J, Ribo M, Vilanova M, et al. (2010) NMR
Spectroscopy Reveals that RNase A is Chiefly Denatured in 40% Acetic Acid,
Implications for Oligomer Formation by 3D Domain Swapping. J Am Chem
Soc 132: 1621–1630.
12. Ercole C, Colamarino RA, Pizzo E, Fogolari F, Spadaccini R, et al. (2009)
Comparison of the structural and functional properties of RNase A and BS-
RNase: a stepwise mutagenesis approach. Biopolymers 91: 1009–1017.
13. Merlino A, Krauss IR, Perillo M, Mattia CA, Ercole C, et al. (2009) Towards an
antitumor form of bovine pancreatic ribonuclease: The crystal structure of three
non-covalent dimeric mutants. Biopolymers.
14. Geiger R, Gautschi M, Thor F, Hayer A, Helenius A (2011) Folding, quality
control, and secretion of pancreatic ribonuclease in live cells. J Biol Chem 286:
5813–5822.
15. D’Alessio G, Riordan J (1997) Ribonucleases: Structures and Functions; press A,
ed. NewYork: Academic press.
16. Tamburrini M, Piccoli R, De Prisco R, Di Donato A, D’Alessio G (1986) Fast
and high-yielding procedures for the isolation of bovine seminal RNAase.
Ital J Biochem 35: 22–32.
17. Berisio R, Sica F, De Lorenzo C, Di Fiore A, Piccoli R, et al. (2003) Crystal
structure of the dimeric unswapped form of bovine seminal ribonuclease. FEBS
Lett 554: 105–110.
18. Mazzarella L, Capasso S, Demasi D, Di Lorenzo G, Mattia CA, et al. (1993)
Bovine seminal ribonuclease: structure at 1.9 A resolution. Acta Crystal-
logr D Biol Crystallogr 49: 389–402.
19. Picone D, Di Fiore A, Ercole C, Franzese M, Sica F, et al. (2005) The role of the
hinge loop in domain swapping. The special case of bovine seminal ribonuclease.
J Biol Chem 280: 13771–13778.
20. Piccoli R, Tamburrini M, Piccialli G, Di Donato A, Parente A, et al. (1992) The
dual-mode quaternary structure of seminal RNase. Proc Natl Acad Sci U S A
89: 1870–1874.
21. Avitabile F, Alfano C, Spadaccini R, Crescenzi O, D’Ursi AM, et al. (2003) The
swapping of terminal arms in ribonucleases: comparison of the solution structure
of monomeric bovine seminal and pancreatic ribonucleases. Biochemistry 42:
8704–8711.
22. Rousseau F, Schymkowitz JW, Itzhaki LS (2003) The unfolding story of three-
dimensional domain swapping. Structure 11: 243–251.
23. Lopez-Alonso JP, Bruix M, Font J, Ribo M, Vilanova M, et al. (2006)
Formation, structure, and dissociation of the ribonuclease S three-dimensional
domain-swapped dimer. J Biol Chem 281: 9400–9406.
24. Ramoni R, Vincent F, Ashcroft AE, Accornero P, Grolli S, et al. (2002) Control
of domain swapping in bovine odorant-binding protein. Biochem J 365:
739–748.
25. Miller KH, Karr JR, Marqusee S (2010) A hinge region cis-proline in
ribonuclease A acts as a conformational gatekeeper for C-terminal domain
swapping. J Mol Biol 400: 567–578.
26. Bergdoll M, Remy MH, Cagnon C, Masson JM, Dumas P (1997) Proline-
dependent oligomerization with arm exchange. Structure 5: 391–401.
27. Ercole C, Spadaccini R, Alfano C, Tancredi T, Picone D (2007) A new mutant
of bovine seminal ribonuclease with a reversed swapping propensity.
Biochemistry 46: 2227–2232.
28. Giancola C, Ercole C, Fotticchia I, Spadaccini R, Pizzo E, et al. (2011)
Structure-cytotoxicity relationships in bovine seminal ribonuclease: new insights
from heat and chemical denaturation studies on variants. Febs J 278: 111–
122.
29. Di Donato A, Galletti P, D’Alessio G (1986) Selective deamidation and
enzymatic methylation of seminal ribonuclease. Biochemistry 25: 8361–8368.
30. Bai Y, Milne JS, Mayne L, Englander SW (1993) Primary structure effects on
peptide group hydrogen exchange. Proteins 17: 75–86.
31. Clarke J, Fersht AR (1996) An evaluation of the use of hydrogen exchange at
equilibrium to probe intermediates on the protein folding pathway. Fold Des 1:
243–254.
32. Bruix M, Ribo M, Benito A, Laurents DV, Rico M, et al. (2008) Destabilizing
mutations alter the hydrogen exchange mechanism in ribonuclease A. Biophys J
94: 2297–2305.
33. Lipari G, Szabo A (1982) Model-free approach to the interpretation of nuclear
magnetic resonance relaxation in macromolecules. 1. Theory and range of
validity. J Am Chem Soc 104: 4546–4559.
34. Alcaraz LA, Del Alamo M, Mateu MG, Neira JL (2008) Structural mobility of
the monomeric C-terminal domain of the HIV-1 capsid protein. Febs J 275:
3299–3311.
35. Daragan VA, Mayo KH (1997) Motional model analyses of protein and peptide
dynamics using 13C and 15N NMR relaxation. Prog Nucl Magn Reson
Spectrosc 31: 63–105.
36. Kroenke CD, Loria JP, Lee LK, Rance M, Palmer AG (1998) Longitudinal and
Transverse 1H–15N Dipolar/15N Chemical Shift Anisotropy Relaxation
Interference: Unambiguous Determination of Rotational Diffusion Tensors
and Chemical Exchange Effects in Biological Macromolecules. J Am Chem Soc
120: 7905–7915.
37. Sessions RB, Gibbs N, Dempsey CE (1998) Hydrogen bonding in helical
polypeptides from molecular dynamics simulations and amide hydrogen
exchange analysis: alamethicin and melittin in methanol. Biophys J 74: 138–152.
38. Dastidar SG, Mukhopadhyay C (2003) Structure, dynamics, and energetics of
water at the surface of a small globular protein: a molecular dynamics
simulation. Phys Rev E Stat Nonlin Soft Matter Phys 68: 021921.
39. Santoro MM, Bolen DW (1992) A test of the linear extrapolation of unfolding
free energy changes over an extended denaturant concentration range.
Biochemistry 31: 4901–4907.
40. Catanzano F, Graziano G, Cafaro V, D’Alessio G, Di Donato A, et al. (1997)
From ribonuclease A toward bovine seminal ribonuclease: a step by step
thermodynamic analysis. Biochemistry 36: 14403–14408.
41. Bernini A, Venditti V, Spiga O, Niccolai N (2009) Probing Protein Surface
Accessibility with Solvent and Paramagnetic Molecules. Progr in NMR Spectr
54: 278–289.
42. Bernini A, Spiga O, Venditti V, Prischi F, Bracci L, et al. (2006) NMR studies of
lysozyme surface accessibility by using different paramagnetic relaxation probes.
J Am Chem Soc 128: 9290–9291.
43. Bennett MJ, Schlunegger MP, Eisenberg D (1995) 3D domain swapping: a
mechanism for oligomer assembly. Protein Sci 4: 2455–2468.
44. Schlunegger MP, Bennett MJ, Eisenberg D (1997) Oligomer formation by 3D
domain swapping: a model for protein assembly and misassembly. Adv Protein
Chem 50: 61–122.
45. Kobe B, Deisenhofer J (1996) Mechanism of ribonuclease inhibition by
ribonuclease inhibitor protein based on the crystal structure of its complex with
ribonuclease A. J Mol Biol 264: 1028–1043.
46. Kover KE, Bruix M, Santoro J, Batta G, Laurents DV, et al. (2008) The solution
structure and dynamics of human pancreatic ribonuclease determined by NMR
spectroscopy provide insight into its remarkable biological activities and
inhibition. J Mol Biol 379: 953–965.
Structure and Dynamics of Monomeric BS-RNase
PLoS ONE | www.plosone.org 10 January 2012 | Volume 7 | Issue 1 | e29076
47. Dechene M, Wink G, Smith M, Swartz P, Mattos C (2009) Multiple solvent
crystal structures of ribonuclease A: an assessment of the method. Proteins 76:861–881.
48. Ercole C, Avitabile F, Del Vecchio P, Crescenzi O, Tancredi T, et al. (2003)
Role of the hinge peptide and the intersubunit interface in the swapping of N-termini in dimeric bovine seminal RNase. Eur J Biochem 270: 4729–4735.
49. Parente A, Branno M, Malorni MC, Welling GW, Libonati M, et al. (1976)Proteolytic enzymes as structural probes for ribonuclease BS-1. Biochim Biophys
Acta 445: 377–385.
50. Richards FM, Vithayathil PJ (1959) The preparation of subtilisn-modifiedribonuclease and the separation of the peptide and protein components. J Biol
Chem 234: 1459–1465.51. Goldschmidt L, Teng PK, Riek R, Eisenberg D (2010) Identifying the amylome,
proteins capable of forming amyloid-like fibrils. Proc Natl Acad Sci U S A 107:3487–3492.
52. D’Alessio G (1999) Evolution of oligomeric proteins. The unusual case of a
dimeric ribonuclease. Eur J Biochem 266: 699–708.53. Delaglio F, Grzesiek S, Vuister GW, Zhu G, Pfeifer J, et al. (1995) NMRPipe: a
multidimensional spectral processing system based on UNIX pipes. J BiomolNMR 6: 277–293.
54. Johnson BA (2004) Using NMRVIEW to visualize and analyze the NMR
spectra of macromolecules. Methods Mol Biol 278: 313–352.55. Sattler M, Schleucher J, Griesinger C (1999) Heteronuclear multidimensional
NMR experiments for the structure determination of proteins in solutionemploying pulsed field gradients. Prog NMR Spectrosc 34: 93–158.
56. Cornilescu G, Delaglio F, Bax A (1999) Protein backbone angle restraints fromsearching a database for chemical shift and sequence homology. J Biomol NMR
13: 289–302.
57. Vuister GW, Delaglio F, Bax A (1993) The use of 1JC alpha H alpha couplingconstants as a probe for protein backbone conformation. J Biomol NMR 3:
67–80.58. Guntert P, Mumenthaler C, Wuthrich K (1997) Torsion angle dynamics for
NMR structure calculation with the new program DYANA. J Mol Biol 273:
283–298.59. Pearlman DA, Case DA, Caldwell JW, Ross WS, Cheatham TEI, et al. (1995)
AMBER, a computer program for applying molecular mechanics, normal modeanalysis, molecular dynamics and free energy calculations to elucidate the
structures and energies of molecules. Comput Phys Commun 91: 1–41.
60. Laskowski RA, Rullmannn JA, MacArthur MW, Kaptein R, Thornton JM
(1996) AQUA and PROCHECK-NMR: programs for checking the quality of
protein structures solved by NMR. J Biomol NMR 8: 477–486.
61. Koradi R, Billeter M, Wuthrich K (1996) MOLMOL: a program for display and
analysis of macromolecular structures. J Mol Graph 14: 51–55, 29–32.
62. Berendsen HJC, van der Spoel D, van Drunen R (1994) GROMACS: A
message-passing parallel molecular dynamics implementation. Computer
Physics Communications 91: 43–56.
63. van Gunsteren WF, Billeter SR, Eising AA, Hunenberger PH, Kruger P, et al.
(1996) Biomolecular Simulation: The GROMOS96 Manual and User Guide.
Zurich: Vdf Hochschulverlag AG an der ETH Zurich.
64. Berendsen HJC, Grigerra JR, Straatsma TP (1987) The missing term in effective
pair potentials. J Phys Chem 91: 6269–6271.
65. Darden T, Perera L, Li L, Pedersen L (1999) New tricks for modelers from the
crystallography toolkit: the particle mesh Ewald algorithm and its use in nucleic
acid simulations. Structure 7: R55–60.
66. Lounnas V, Pettitt BM (1994) A connected-cluster of hydration around
myoglobin: correlation between molecular dynamics simulations and experi-
ment. Proteins 18: 133–147.
67. De Simone A, Dodson GG, Verma CS, Zagari A, Fraternali F (2005) Prion and
water: tight and dynamical hydration sites have a key role in structural stability.
Proc Natl Acad Sci U S A 102: 7535–7540.
68. Tellinghuisen J (2000) Nonlinear Least-Squares Using Microcomputer Data
Analysis Programs: KaleidaGraphTM in the Physical Chemistry Teaching
Laboratory. J Chem Educ 77: 1233.
69. Farrow NA, Muhandiram R, Singer AU, Pascal SM, Kay CM, et al. (1994)
Backbone dynamics of a free and phosphopeptide-complexed Src homology 2
domain studied by 15N NMR relaxation. Biochemistry 33: 5984–6003.
70. Tessari M, Vis H, Boelens R, Kaptein R, Vuister GW (1997) Quantitative
Measurement of Relaxation Interference Effects between 1HN CSA and 1H–
15N Dipolar Interaction: Correlation with Secondary Structure. J Am Chem
Soc 119: 8985–8990.
71. Goldman M (1984) Interference Effects in the Relaxation of a Pair of Unlike
Spin-1/2 Nuclei. Journal of Magnetic Resonance 60: 437–452.
72. Varrazzo D, Bernini A, Spiga O, Ciutti A, Chiellini S, et al. (2005) Three-
dimensional computation of atom depth in complex molecular structures.
Bioinformatics 21: 2856–2860.
Structure and Dynamics of Monomeric BS-RNase
PLoS ONE | www.plosone.org 11 January 2012 | Volume 7 | Issue 1 | e29076